structural, biochemical, and physiological characterization of photosynthesis in two c4 subspecies...

Journal of Experimental Botany, Vol. 59, No. 7, pp. 1715–1734, 2008

doi:10.1093/jxb/ern028 Advance Access publication 26 March, 2008

SPECIAL ISSUE RESEARCH PAPER

Structural, biochemical, and physiological characterizationof photosynthesis in two C4 subspecies of Tecticorniaindica and the C3 species Tecticornia pergranulata(Chenopodiaceae)

Elena V. Voznesenskaya1, Hossein Akhani2, Nuria K. Koteyeva1, Simon D. X. Chuong3, Eric H. Roalson4,

Olavi Kiirats4, Vincent R. Franceschi4 and Gerald E. Edwards4,*

1 Laboratory of Anatomy and Morphology, V. L. Komarov Botanical Institute of Russian Academy of Sciences,Prof. Popov Street 2, 197376, St Petersburg, Russia2 Department of Plant Sciences, School of Biology, College of Sciences, University of Tehran,PO Box 14155-6455, Tehran, Iran3 Department of Biology, University of Waterloo, Waterloo, Ontario N2L 3G1, Canada4 School of Biological Sciences, Washington State University, Pullman, WA 99164-4236, USA

Received 4 September 2007; Revised 16 January 2008; Accepted 21 January 2008

Abstract

Among dicotyledon families, Chenopodiaceae has the

most C4 species and the greatest diversity in structural

forms of C4. In subfamily Salicornioideae, C4 photo-

synthesis has, so far, only been found in the genus

Halosarcia which is now included in the broadly

circumscribed Tecticornia. Comparative anatomical,

cytochemical, and physiological studies on these taxa,

which have near-aphyllous photosynthetic shoots,

show that T. pergranulata is C3, and that two sub-

species of T. indica (bidens and indica) are C4 (Kranz-

tecticornoid type). In T. pergranulata, the stems have

two layers of chlorenchyma cells surrounding the

centrally located water storage tissue. The two sub-

species of T. indica have Kranz anatomy in reduced

leaves and in the fleshy stem cortex. They are

NAD-malic enzyme-type C4 species, with mesophyll

chloroplasts having reduced grana, characteristic of

this subtype. The Kranz-tecticornoid-type anatomy is

unique among C4 types in the family in having groups

of chlorenchymatous cells separated by a network of

large colourless cells (which may provide mechanical

support or optimize the distribution of radiation in the

tissue), and in having peripheral vascular bundles with

the phloem side facing the bundle sheath cells. Also,

the bundle sheath cells have chloroplasts in a centrifu-

gal position, which is atypical for C4 dicots. Fluores-

cence analyses in fresh sections indicate that all

non-lignified cell walls have ferulic acid, a cell wall

cross-linker. Structural–functional relationships of C4

photosynthesis in T. indica are discussed. Recent

molecular studies show that the C4 taxa in Tecticornia

form a monophyletic group, with incorporation of

the Australian endemic genera of Salicornioideae, in-

cluding Halosarcia, Pachycornia, Sclerostegia, and

Tegicornia, into Tecticornia.

Key words: C3 plants, C4 plants, Chenopodiaceae, chloroplast

ultrastructure, Halosarcia, immunolocalization, NAD-ME type,

photosynthetic enzymes, phylogeny, Tecticornia.

Introduction

In the family Chenopodiaceae, which has C3 and C4

species, all C4 genera occur in subfamily Chenopodioid-eae (Atriplex) and in a succulent clade made up of three

* To whom correspondence should be addressed. E-mail: [email protected]: BS, bundle sheath; BSC, bundle sheath cell; Ci, intercellular levels of CO2; CW, cell wall; C*, CO2 compensation point based on Rubiscocarboxylase/oxygenase activity; GDC, glycine decarboxylase; MC, mesophyll cell; ML, maximum likelihood; NAD-ME, NAD-malic enzyme; NADP-ME,NADP-malic enzyme; PEPC, phophoenolpyruvate carboxylase; PEP-CK, phosphoenolpyruvate carboxykinase; PPDK, pyruvate; Pi, dikinase; PPFD,photosynthetic photon flux density; WS, water storage.

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subfamilies: Suaedoideae (Suaeda and Bienertia),Salsoloideae (various genera), and Salicornioideae(Halosarcia) (Carolin et al., 1975; Pyankov, 1991;Akhani et al., 1997; Jacobs, 2001; Pyankov et al., 2001a;Kadereit et al., 2003; Kapralov et al., 2006; Akhaniand Ghasemkhani, 2007). This family has the largestnumber of C4 species and also the greatest diversity inleaf anatomy among dicot families, including C4 Kranzand C4 single-cell type species, as well as C3 type species(Carolin et al., 1975; Sage et al., 1999; Edwards et al.,2004). Six C4 types of Kranz anatomy (atriplicoid,kochioid, salsoloid, halosarcoid, and, in the genusSuaeda, salsina and schoberia types) and five C3 types(axyroid, corispermoid, austrobassioid, neokochioid,and sympegmoid) have been described among speciesof this family, mostly in corresponding genera(Carolin et al., 1975, 1982; Voznesenskaya, 1976b;Voznesenskaya and Gamaley, 1986; Jacobs, 2001;Kadereit et al., 2003). Recently, leaf anatomy in represen-tative Chenopodiaceae species was further revised withthe description of 15 C4 types (Kadereit et al., 2003).The C4 types of anatomy vary in the structure andarrangement of the two-layered chlorenchyma adjacentto the vascular bundles, and by the presence or absenceof water storage (WS) tissue, hypodermal cells, andsclerenchyma, and whether they have continuous orinterrupted Kranz tissue.Species in subfamily Salicornioideae are hygrohalo-

phytic plants which belong to the most salt-tolerantangiosperms inhabiting salt marshes and inland salinehabitats. In this subfamily, only one species, Halosarciaindica, has been identified as C4 on the basis of itsanatomy and C4-type carbon isotope composition, while11 species of this genus have C3-type carbon isotopecomposition (Wilson, 1980; Carolin et al., 1982; Akhaniet al., 1997).Carolin et al. (1982) studied the anatomical structure in

several representatives of the genus Halosarcia. Species withC3-type carbon isotope values had 2–3 layers of chloren-chyma tissue surrounding WS parenchyma, while severalsubspecies of H. indica had C4-type isotope values andKranz anatomy. Unlike the salsoloid type of Kranz anatomy,an unusual occurrence of colourless (or, more accurately,organelle-deficient) cells between groups of chlorophyllousmesophyll cells (MCs) was reported in H. indica.

Halosarcia was segregated from Arthrocnemum byWilson (1980) by the absence of sclereids in chloren-chyma tissue and by the flowers having a single stamen.According to recent phylogenies, Halosarcia is placed ina monophyletic clade with four other Australian endemicgenera, including Tecticornia, Pachycornia, Sclerostegia,and Tegicornia (Shepherd et al., 2004; Kadereit et al.,2006). Shepherd and Wilson (2007) have incorporated allthese genera into a broadly defined Tecticornia s. l. whichis accepted in this paper.

The aim of the present study was to characterize theanatomy and ultrastructure of chlorenchyma, and theunusual occurrence of colourless cells within Kranzanatomy, to identify the C4 biochemical subtype, andanalyse features of CO2 fixation in T. indica (using twosubspecies which occur on different continents and arevisibly different, bidens and indica). Comparative analy-ses were made with the C3 species T. pergranulata. Thephylogenetic position of these representatives of Tecticor-nia in subfamily Salicornioideae was also evaluated.

Materials and methods

Plant material

Seeds of T. pergranulata (J. M. Black) K. A. Sheph. & PaulG. Wilson subsp. pergranulata and T. indica subsp. bidens (Nees)K. A. Sheph. & Paul G. Wilson were provided by G Barrett, GregBarrett & Associates, Darlington, Western Australia. Seeds ofT. indica (Willd.) K. A. Sheph. & Paul G. Wilson subsp. indicawere collected by H Akhani from Pakistan, 40 km NW of Karachi(H. Akhani 16537). Seeds were stored at 3–5 �C prior to use, thengerminated on moist paper in Petri dishes in a growth chamber at30/25 �C and a photosynthetic photon flux density (PPFD) of75 lmol m�2 s�1 with a 14/10 h light/dark photoperiod. Theseedlings were then transplanted to 10 cm diameter pots withcommercial potting soil and grown for 3 d under the same regime.Established plants were then transferred to a growth chamber(model GC-16; Enconair Ecological Chambers Inc., Winnipeg,Canada) and grown under ;400 PPFD with a 16/8 h light/darkphotoperiod and 25/18 �C day/night temperature regime. Formicroscopy and biochemical analyses, samples of mature segmentswere taken from ;2.5- to 3-month-old plants.Voucher specimens are available at the Marion Ownbey

Herbarium, Washington State University: T. pergranulata(E. Voznesenskaya 28), April 2006, WS 369799; T. indica subsp.indica (E. Voznesenskaya 29), April 2006, WS 369801; T. indicasubsp. bidens (E. Voznesenskaya 27), April 2006, WS 369800.

Light and electron microscopy

Hand-cut sections of fresh stems were placed in water and studiedunder a light stereo microscope. The area of chlorenchyma tissueexternal to the central cylinder, and of WS tissue, as a percentage ofthe total cross-sectional area was determined from digital images(on ;10 cross-sections taken from two different plants) usingUTHSCSA, Image Tool for Windows, version 3.00, University ofTexas Health Science Center, San Antonio, TX, USA.For microscopy on fixed material, samples were taken from 2–3

plants (5–6 samples from 2–3 branches of each plant). Samples forstructural studies were fixed at 4 �C in 2% (v/v) paraformaldehydeand 2% (v/v) glutaraldehyde in 0.1 M phosphate buffer (pH 7.2),post-fixed in 2% (w/v) OsO4, and then, after a standard acetonedehydration procedure, embedded in Spurr’s epoxy resin. Cross-sections were made on a Reichert Ultracut R ultramicrotome(Reichert-Jung GmbH, Heidelberg, Germany). For light micros-copy, semi-thin sections were stained with 1% (w/v) toluidine blueO in 1% (w/v) Na2B4O7. Ultra-thin sections were stained fortransmission electron microscopy with 2% (w/v) uranyl acetatefollowed by 2% (w/v) lead citrate. Hitachi H-600 (Hitachi ScientificInstruments, Mountain View, CA, USA) and JEOL JEM-1200 EX(JEOL USA, Inc., Peabody, MA, USA) transmission electronmicroscopes were used for observation and photography.

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For scanning electron microscopy (SEM), leaf samples werefixed at 4 �C in 2% (v/v) paraformaldehyde and 2% (v/v)glutaraldehyde in 0.1 M phosphate buffer (pH 7.2), post-fixed in2% (w/v) OsO4, and then dehydrated in an ethanol series to 100%ethanol, cryofractured in liquid nitrogen, critical-point dried,attached to SEM mounts, sputter-coated with gold, and observedwith a Hitachi S570 SEM (Hitachi, Ltd, Tokyo, Japan).The size of chloroplasts and mitochondria, and the thickness of

cell walls (CWs), were measured on micrographs from leaf cross-sections with an image analysis program (Image Tool forWindows). For measurements of the length and width, images ofchloroplast median sections were used. For determining the size ofmitochondria, the small diameter of profiles on cross-sections wasmeasured. As was previously noted in quantitative studies onmitochondria, rather long profiles can occasionally be observed inmicroscopy sections; however, only the small diameter will reflectthe difference in size between different tissues or species (seeVoznesenskaya et al., 2007).

Fluorescence of chloroplasts and cell walls, and lignification

Hand-cut sections of leaves or stems were placed on slides indistilled water and examined under UV light [with a 4#,6-diamidino-2-phenylindole (DAPI) filter] with a Zeiss LSM 510META (Jena, Germany) microscope. For comparison, similarsections were treated with 0.1 M NH4OH to reveal the presence ofCW-bound ferulic acid. According to Harris and Hartley (1976,1980), Hartley and Harris (1981), and Rudall and Caddick (1994),if the tissue contains CW-bound ferulic acid, an increase of the pH(to ;10.3) will change the blue fluorescence of CWs to blue-greenby ionization of the phenol OH group. This treatment does notchange the autofluorescence of CWs in lignified or suberizedtissues. To detect the position of lignified tissue, sections weretreated for 1 h with phloroglucinol (2% in 10% HCl), which stainslignin-containing CWs red, while for detection of suberization,sections were stained with Sudan IV in 70% alcohol, which stainssuberized CWs dark red (Ruzin, 1999).

In situ immunolocalization

Leaf samples were fixed at 4 �C in 2% (v/v) paraformaldehyde and1.25% (v/v) glutaraldehyde in 0.05 M PIPES buffer, pH 7.2. Thesamples were dehydrated with a graded ethanol series andembedded in London Resin White (LR White, Electron MicroscopySciences, Fort Washington, PA, USA) acrylic resin. Antibodiesused (all raised in rabbit) were anti-Spinacia oleracea L. Rubisco(LSU) IgG (courtesy of B McFadden), commercially available anti-Zea mays L. phosphoenolpyruvate carboxylase (PEPC) IgG(Chemicon, Temecula, CA, USA), anti-pyruvate, Pi dikinase(PPDK) IgG (courtesy of T Sugiyama), anti-Amaranthus hypochon-driacus L. mitochondrial NAD-malic enzyme (NAD-ME) IgG(courtesy of J Berry), which was prepared against the 65 kDaa-subunit (Long and Berry, 1996), and anti-Pisum sativumL. glycine decarboxylase (GDC) against the P subunit (courtesy ofD Oliver). Pre-immune serum was used in all cases for controls.Cross-sections, 0.8–1 lm thick, were dried from a drop of water

onto gelatin-coated slides and blocked for 1 h with TBST+BSA[10 mM TRIS-HCl, 150 mM NaCl, 0.3% (v/v) Tween-20, 1% (w/v)bovine serum albumin, pH 7.2]. They were then incubated for 3 hwith either pre-immune serum diluted in TBST+BSA (1:100), anti-Rubisco LSU (1:500), or anti-PEPC (1:200). The slides werewashed with TBST+BSA and then treated for 1 h with proteinA–gold (10 nm particles diluted 1:100 with TBST+BSA). Afterwashing, the sections were exposed to a silver enhancement reagentfor 20 min according to the manufacturer’s directions (Amersham,Arlington Heights, IL, USA), stained with 0.5% (w/v) Safranin O,

and imaged in a reflected/transmitted mode using a BioRad 1024confocal system with a Nikon Eclipse TE 300 inverted microscopeand Lasergraph image program 3.10. The background labelling withpre-immune serum was very low, although some infrequentlabelling occurred in areas where the sections were wrinkled due totrapping of antibodies/label (results not shown).For TEM immunolabelling, thin sections on Formvar-coated

nickel grids were incubated for 1 h in TBST+BSA to block non-specific protein binding on the sections. They were then incubatedfor 3 h with either the pre-immune serum diluted in TBST+BSA(1:50) or anti-PEPC (1:20), anti-Rubisco (1:50), anti-PPDK (1:40),anti-NAD-ME (1:50), or anti-GDC (1:50) antibodies. After washingwith TBST+BSA, the sections were incubated for 1 h with proteinA–gold (10 or 15 nm) diluted 1:100 with TBST+BSA. The sectionswere washed sequentially with TBST+BSA, TBST, and distilledwater, and then post-stained with a 1:4 dilution of 1% (w/v)potassium permanganate and 2% (w/v) uranyl acetate. Images werecollected using a JEOL JEM-1200 EX transmission electronmicroscope. The density of labelling was determined by countingthe gold particles on electron micrographs and calculating thenumber per unit area (lm2).

Staining for polysaccharides

The periodic acid–Schiff’s procedure (PAS) was used for stainingstarch in sectioned materials. Sections, 0.8–1 lm thick, were madefrom the same samples used for immunolocalization, dried ontogelatin-coated slides, incubated in periodic acid [1% (w/v)] for30 min, washed, and then incubated with Schiff’s reagent (Sigma,St Louis, MO, USA) for 1 h. After rinsing, the sections were readyfor analysis by light microscopy. CWs and starch stained brightreddish pink, while other elements of the cells (cytoplasm) remainedunstained. Controls lacking the periodic acid treatment (required foroxidation of the polysaccharides giving rise to Schiff’s-reactivegroups) showed little or no background staining (not shown).

Western blot analysis

Total proteins were extracted from leaves by homogenizing 500 mgof tissue in 1 ml of extraction buffer [100 mM TRIS-HCl, pH 7.5,5 mM MgSO4, 10 mM dithiothreitol, 5 mM EDTA, 0.5% (w/v)SDS, 2 % (v/v) b-mercaptoethanol, 10% (v/v) glycerol, 1 mMphenylmethylsulphonyl fluoride, and 25 lg ml�1 each of aprotinin,leupeptin, and pepstatin]. After centrifugation at high speed for3 min in a microcentrifuge, the supernatant was collected and theprotein concentration was determined by Bradford protein assay(Bio-Rad) using BSA as a standard. Protein samples (10 lg) wereseparated by 12.5% SDS–PAGE, blotted onto nitrocellulose, andprobed with anti-A. hypochondriacus NAD-ME (1:5000), anti-Z.mays NADP-malic enzyme (NADP-ME), courtesy of C Andreo(Maurino et al., 1996) (1:5000), anti-Z. mays PEPC (1:10 000),anti-Z. mays PPDK (1:5000), anti-Urochloa maxima phosphoenol-pyruvate carboxykinase (PEP-CK), courtesy of RC Leegood, oranti-Spinacia oleracea Rubisco LSU (1:10 000) overnight at 4 �C.Goat anti-rabbit IgG–alkaline phosphatase conjugate antibody (Bio-Rad) was used at a dilution of 1:50 000 for detection. Boundantibodies were localized by developing the blots with 20 mMnitroblue tetrazolium and 75 mM 5-bromo-4-chloro-3-indolyl phos-phate in the detection buffer (100 mM TRIS-HCl, pH 9.5, 100 mMNaCl, and 5 mM MgCl2).

Acidity

Plant samples were collected just before the beginning of the lightperiod, in the middle of the day, and in the late afternoon just beforethe beginning of the dark period. Samples of known fresh weight(between 0.2 g and 0.5 g) were ground in 2 ml of distilled water.

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The sample was titrated with 0.01 M NaOH to a pH 7 end pointusing a pH meter, and the leq acid per g fresh weight wascalculated.

Measurements of rates of photosynthesis

Rates of photosynthesis in response to light were measured witha CO2 analyser (ADC LCPro+, ADC BioScientific Ltd, Hoddesdon,UK) operating in a differential mode. The air temperature was2560.5 �C (stem temperature was 25–27 �C), the minimumhumidity was 12.060.5 mbar, and the flow rate was 200 lmol s�1.The local average barometric pressure, as determined by the CO2

analysing system, was 922.363.4 mbar.For each experiment, part of a branch of an intact plant (3–4 months

old) was enclosed in the conifer chamber designed for terete orsemi-terete leaves. The branch was illuminated with 920 PPFDunder 370 lbar CO2 until a steady-state rate of CO2 fixation wasobtained (generally 40–50 min). For varying light experiments at370 lbar CO2, measurements were made beginning at 1380 PPFD,followed by decreasing increments of light intensity at 4 minintervals.For measurement of the response of photosynthesis to varying

CO2 at 2% and 21% O2, and for determining the CO2 compensationpoint based on Rubisco carboxylase/oxygenase activity (C*), gasexchange was measured with the FastEst gas system (see Laisk andEdwards, 1997; Sun et al., 1999). A branch was enclosed in a smallleaf chamber (4 cm33 cm30.5 cm) with an open gas flow rate of0.5 mmol s�1. The chamber temperature was maintained at 25 �C,with the water jacket of the chamber connected to a thermostatedwater bath. Both sides of the branch were illuminated with a PPFDof 900 lmol quanta m�2 s�1 (measured with a Li-Cor 185 quantumsensor) at the glass window by fibreoptics with a Schott KL1500source (H Walz, Effeltrich, Germany). Relative humidity in the leafchamber was controlled by diverting part of the air flow streamthrough air that was equilibrated with water at 50 �C. CO2 and O2

partial pressures were obtained by mixing pure CO2, O2, N2, andCO2-free air with the help of capillaries. The pressure difference inthe capillaries was stabilized by manostats (tubes with open endssubmerged in water to adjustable heights). The water vapourpressure was measured with a psychrometer. CO2 exchange wasmeasured with a MK3-225 IR gas analyser (ADC, Hoddesdon,Hertfordshire, UK) or a Li-6251 analyser (Li-Cor, Lincoln, NE,USA). Data were recorded by computer using an A/D board ME-30and a RECO program, and analysed by computer programs ANALand SYNTE. The programs RECO and ANAL were written by VOva (University of Tartu, Estonia) in Turbo-Pascal. The intercellu-lar CO2 concentration in the leaf was calculated with inputs for therate of photosynthesis, the CO2 concentration in the air, and thediffusive resistance of CO2 from the atmosphere to the intercellularspace. The latter was calculated by determining the diffusiveresistance to water by measuring transpiration, and the water vapourconcentration difference from the leaf to air (for a description seeKu et al., 1977; von Caemmerer and Farquhar, 1981). The C*,where the rate of CO2 uptake equals photorespiratory loss of CO2,was determined by taking the co-ordinates of the intersection ofCO2 response curves measured at different light intensities (Brooksand Farquhar, 1985).The area of tissue exposed to incident light was calculated by

taking a digital image of the branch that was enclosed in thechamber, and then determining the exposed branch area using animage analysis program (Image Tool for Windows).

d13C values

Measures of the carbon isotope composition (d13C values) weremade at Washington State University on leaf and stem samples

taken from plants using a standard procedure relative to PDB (PeeDee Belemnite) limestone as the carbon isotope standard (Benderet al., 1973). Plant samples (from plants growing in the WashingtonState University School of Biological Sciences growth chamber)were dried at 80 �C for 24 h, milled to a fine powder, and then1–2 mg were placed in a tin capsule and combusted in a Eurovectorelemental analyser. The resulting N2 and CO2 gases were separatedby gas chromatography and admitted into the inlet of a MicromassIsoprime isotope ratio mass spectrometer (IRMS) for determinationof 13C/12C ratios (R). d13C values were determined whered¼10003(Rsample/Rstandard)�1.

Statistics

Where indicated, standard errors were determined, and analysis ofvariance (ANOVA) was performed with Statistica 7.0 software(StatSoft, Inc.). Tukey’s HSD (honest significant difference) testswere used to analyse differences between cell types. All analyseswere performed at the 95% significance level.

Results

General features including the stem surface

Plants of all three representatives are prostrate to erectshrubs and subshrubs with stems comprised of segmentswith intercalary growth. These plants have reducedopposite leaves (;1 mm in length) at the distal (top) endof each segment (Fig. 1B, G, L). Photosynthesis isaccomplished in the fleshy cortex of the articulated shoots.Under the growth conditions used, T. pergranulata(Fig. 1A) was fast growing, having bright-green stemswhich were 2–3 mm in diameter (Fig. 1B), T. indicasubsp. indica (from Pakistan) had thicker stems (diameter4–5 mm) with dark- or purple-green colour (Fig. 1F),while T. indica subsp. bidens (from Australia) had thinstems with a bright-green colour (Fig. 1J), resemblingT. pergranulata. In T. indica subsp. indica, the segmentsin the vegetative branches are compact with formation ofa cylindrical jointed stem (Fig. 1F), in contrast to T. indicasubsp. bidens, whose stems are longer and narrowertowards the base, resulting in a moniliform jointed stem(Fig. 1K). Figure 1E shows plants of T. indica subsp.indica in a natural habitat in Pakistan.All three taxa have morphology which is typical for

members of subfamily Salicornioideae, including shortinternodes and nearly aphyllous shoots with scale-likeleaves (Fig. 1). The cylindrical stem has a fleshy cortexwith chlorenchyma on the periphery, which is characteris-tic of all species in the subfamily. Sunken anomocyticstomata are mostly distributed in vertical rows on theepidermis of the fleshy cortex of the segments, alternat-ing with rows of cells without stomata, with their longaxis oriented perpendicular to the axis of the stem(Fig. 1C, H, L, M, light bands, and D, I). In all species,stomata are located throughout the epidermis of the fleshycortex of the segment and leaf, but they are absent in thetransparent leaf marginal area and on the abaxial




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epidermis along the leaf main rib (Fig. 1F, G, K, L). In C4

T. indica, stomata are located only in the epidermal cellswhich are external to the groups of chlorenchyma cells(Fig. 2K).

Light microscopy

The stem tissue of T. pergranulata has C3 anatomy, withtwo layers of mesophyll chlorenchyma surrounding theperiphery of the cortex with WS tissue in the centre(Fig. 2A–D). In the reduced leaves of T. pergranulata, thechlorenchyma tissue occurs only on the abaxial side(results not shown, but similar to that of Salicorniafruticosa; see Fahn and Arzee, 1959).In the stems of T. pergranulata, most of the peripheral

vascular bundles are located in WS tissue one cell apartfrom the chlorenchyma cells, and they are distributed withthe phloem facing towards the chlorenchyma, with thecentral cylinder in the centre of the stem. There are largeintercellular air spaces beneath the stomata (also seeCarolin et al., 1982). In this species, chlorenchyma tissuecomprises ;35% and WS tissue ;60% of the total area of

stem cross-section. Starch grains are abundant throughoutall chlorenchyma cells, with the highest density in theoutermost layer (Fig. 2D).In both subspecies of T. indica, analysis of cross-

sections of the young shoot segments showed that themain volume of fleshy cortex is comprised of WStissue (Fig. 2E, L). In subspecies indica (Fig. 2E), theperipheral chlorenchyma tissue is 15–20% while theWS parenchyma is 70–75% of the total area of the stemcross-section. In T. indica subsp. bidens (Fig. 2L), thestems are thinner, and the tissue in the chlorenchymatousrings is 20–30% of the total area of the stem segment(depending on the position of the section from thenode). As in T. pergranulata, chlorenchyma tissueoccurs only on the abaxial side of the reduced leavesin both subspecies of T. indica (Fig. 2M). In the stems ofT. indica, small peripheral vascular bundles are distributeddirectly under the bundle sheath cells (BSCs).Both subspecies of T. indica have two layers of

chlorenchyma, which are characteristic of C4 species withKranz anatomy, an outer layer of palisade MCs and an

Fig. 1. General view of plants of Tecticornia pergranulata (A–D) and T. indica (E–I, subsp. indica and J–M, subsp. bidens), and characteristics ofstems and their surfaces. Except for E, all images are from plants grown in WSU growth chambers. Tecticornia pergranulata: (A) plant ;3 monthsold. (B) Branch. (C) Stem surface. Light bands show the position of stomata. (D) Stem surface (SEM), showing the position of stomata. Tecticorniaindica subsp. indica: (E) natural habitat. (F) Plant ;3 months old; the insert is the tip of a branch. (G) Leaf, view from the top (apical part up, abaxialside to the right). (H) Stem surface with light bands showing the position of stomata. (I) Stem surface, SEM. Tecticornia indica subsp. bidens:(J) plant ;3 months old. (K) Branch. (L) Tip of the branch. (M) Stem surface. Light bands show the position of stomata. S, stomata. Scale bars: 1 cmfor A; 3 mm for B, F (inset), K; 5 cm for F, J; 1 mm for G; 0.5 mm for C, L, M; 100 lm for D, H, I.




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inner layer of BSCs (Fig. 2E–G, J, L). The unusual featureof these species is the presence of large colourless MCsseparating groups of chlorenchymatous palisade MCs(Fig. 2F–I, L). Observations of paradermal sections showthat the islands of chlorenchyma cells are surrounded bya network of large colourless MCs which consist of 1–3cells across (Fig. 2H, I). In hand-cut paradermal sections,it was also noticed that there is no green colour in BSCsin some regions where there are colourless MCs betweenthe chlorenchyma cells (Fig. 2H). More careful studiesshowed that there are sparse, nearly empty cells in thelayer of BSCs which are located under colourless MCs

(Fig. 2J); colourless BSCs were observed more often inT. indica subsp. indica. The colourless BSCs locatedunder groups of colourless MCs appear to have no, orlimited, contact with the neighbouring chlorenchymatousMCs. There are rather large intercellular air spacesbetween the epidermal and chlorenchyma cells beneaththe stomata (substomatal cavity), while between stomatathe MCs are closely associated with epidermal cells(Fig. 2J, K). Larger intercellular air spaces also occurbetween the distal ends of MCs in both subspecies ofT. indica; whereas, at the proximal ends, all MCs are closeto each other with little or no intercellular air space,

Fig. 2. Hand-cut sections (A, E, H, L), SEM (B, F), general anatomy (C, G, I, J, M–P), and periodic acid–Schiff’s (PAS) staining procedure forcarbohydrates (D, K) of stems of T. pergranulata (A–D) and T. indica (E–K subsp. indica and L–P subsp. bidens). (A–G, J–L) Cross-sections of thestems. (D, K) PAS staining for carbohydrates showing starch localization. (H) Hand-cut section and (I) resin-embedded paradermal section of thestem showing the distribution of the chlorenchyma and colourless cells. (M) Longitudinal section of the tip of the shoot showing the positioning ofthe chlorenchyma only along the abaxial side of the leaf and its absence on the adaxial side and along the leaf tip. (N–P) Longitudinal sections ofa young segment showing the development of chlorenchyma from the base (N) through the middle (O) to the tip of the leaf (P). BS, bundle sheathcells; C, chlorenchyma; CBS, colourless bundle sheath cells; CC, central cylinder; CM, colourless mesophyll; M, mesophyll cells; SC, substomatalcavity; WS, water storage tissue. Scale bars: 1 mm for A, E; 100 lm for B–D, H, N–P; 50 lm for F, G, I, J, K; 0.5 mm for L, M.




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depending on the subspecies studied: in subsp. bidensthere are more intercellular air spaces where the bundlesheath (BS) CW faces the MCs, and, in general, cells aremore tightly packed in subsp. indica. Starch granules areabundant in BSC chloroplasts of both subspecies (resultsshown only for subsp. indica, Fig. 2K).Development of the two-layered chlorenchyma tissue in

T. indica subsp. bidens is shown in the longitudinalsection of the shoot tip (Fig. 2M) and the young segment(Fig. 2N–P). Both layers of photosynthetic cells evidentlyoriginate from one layer of pre-chlorenchyma cells duringleaf development (Fig. 2M) and during formation of thecortex chlorenchyma in the internodal meristem (Fig. 2N).In the outer row of chlorenchyma, there are cells withdifferent levels of development, with some having a lowercytoplasmic content which could be distinguished ata rather early stage (Fig. 2O, P). Presumably, these cellswith lower cytoplasmic content are precursors to theformation of the colourless MCs.

Fluorescence of chloroplasts and cell walls, andlignification

For all three representative taxa, fresh hand-cut sectionsplaced in water have red fluorescence from chloroplasts inthe outer chlorenchyma layers, with lower intensity redfluorescence coming from the pith, and from parenchymatissue between the central cylinder and a suberized layer(which is considered periderm, e.g. see Discussion, andArcihovskii, 1928; Vosnesenskaya and Steshenko, 1974).In all three taxa there was very bright blue fluorescence ofCWs under UV light (Fig. 3A, D, G). Since it is knownthat lignified and suberized CWs have bright bluefluorescence, the sections were treated with phloroglucinolto test for lignification; the results are shown in Fig. 3B,E, H. Staining with phloroglucinol changed the colour ofxylem, sclerenchymatous tissue, and mechanical extraxyl-ary fibres to dark red, showing the presence of lignifica-tion only in these tissues (Fig. 3B, E, H). The bluefluorescing CWs of WS tissue did not change their colour.Several cell layers outside the central cylinder, havingespecially bright light-blue fluorescence in sections placedin water, changed their colour slightly to red withphloroglucinol treatment. Staining of sections with SudanIV changed the colour of CWs outside the central cylinderto dark red, showing the presence of suberin (not shown).Thus, the blue fluorescence of CWs of WS and other cellsis not related to lignification or suberization in thesespecies.Sections were then treated with NH4OH to check for the

presence of bound ferulic acid. In all three representatives,under alkaline conditions the blue fluorescence of all non-lignified CWs became more intense and changed colour togreen, demonstrating the presence of CW-bound ferulicacid. In contrast, the colour of CWs of all xylem vessels

in the central cylinder and in small vascular bundles,sclerenchymatous tissue in the central cylinder, and raremechanical fibres outside the suberized layer remainedbright blue under alkaline conditions (Fig. 3C, F, I),indicative of lignified or suberized CWs.In all three Tecticornia representatives, the most in-

tensive blue fluorescence of CWs in sections in water wasin the epidermis, WS tissue, the 2–3 layers of thick-walledperidermal cells outside the central cylinder, and mechan-ical tissues surrounding vascular bundles, together withthe xylem (Fig. 3A, D, G). Furthermore, in both C4

subspecies, BS and colourless mesophyll CWs fluorescemore intensively than chlorenchymatous mesophyll CWs.Morphometrical study of CW thickness showed that allthree taxa have a rather thick outer epidermal CW, whichwas thickest in T. indica subsp. indica (Table 1).Chlorenchyma MCs have thin CWs (;0.07–0.08 lm) inall three representatives. BSCs of both T. indica sub-species have rather thick CWs (;0.8 lm in subsp. indicaand ;0.5 lm in subsp. bidens). The thickness of CWs inWS tissue and colourless MCs is similar to the thicknessof BS CWs for both T. indica subspecies, with greaterthickness in subsp. indica. The thickness of the CW inWS tissue in T. pergranulata is also more than twice thatof the mesophyll CW, but much lower than in the twosubspecies of T. indica (Table 1). Thus, the higherfluorescence in the CW of the outer epidermal, BS, andWS tissue appears related to the greater thickness of CWin these tissues.

Electron microscopy

Stems of all three Tecticornia taxa are covered by a thickcuticle which has a structure typical of many desertchenopods, with a rather well-formed outer lamellatedlayer of cuticle proper, followed by the inner cuticularlayer with intensive development of reticulated poly-saccharide microfibrils, also called dendrites (Fig. 4A, E).The thickness of the cuticle layer depends on the ageof the segment, but, in general, the thickest cuticle wasfound in T. indica subsp. indica (;2 lm) while theother two taxa, T. pergranulata and T. indica subsp.bidens, have rather similar cuticle thickness of ;0.7 lm(Table 1). The thickness of the outer epidermal CWvaries from 1.1 lm in T. indica subsp. bidens, to 1.7 lmin T. pergranulata, to 3 lm in T. indica subsp. indica(Table 1).In chlorenchyma cells of T. pergranulata, the chloro-

plasts, which are located mostly towards the intercellularspaces, have grana consisting of 8–10 thylakoids (Fig.4B). Mitochondria are rather small (;0.4 lm, Table 2),and have falciform cristae, which is typical for C3 species.MCs have a thin CW, 0.08 lm (measured between twoadjacent MCs divided by 2, Table 1), which is similar tothat measured at the intercellular air space (not shown).




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Fig. 3. Blue-green fluorescence under UV light and lignification in the hand-cut cross-sections of T. pergranulata (A–F) and T. indica subsp. indica(G–I). (A, D, G) Blue autofluorescence of the CW in fresh sections placed in water. (B, E, H) Staining with phloroglucinol changes the colour ofmechanical tissues and xylem to dark red, showing the presence of lignified CWs. (C, F, I) Light-green fluorescence of CWs in sections placed in0.1 M NH4OH. C, chlorenchyma; CC, central cylinder; M, mesophyll cells; SL, suberized layer; WS, water storage tissue. Scale bars: 350 lm for Aand C; 500 lm for B and H; 150 lm for D–F; 200 lm for G and I.

Table 1. Thickness of the cuticle and cell walls in Tecticornia species (lm)

Analyses were made by one-way ANOVA with Tukey’s HSD. Means followed by a different lower-case letter within a row indicate a significantdifference between cell types (P <0.05). Means followed by a different upper-case letter within a column indicate a significant difference betweenspecies (P <0.05).

Species Cell wall

Cuticle Outerepidermal

Chlorophyllousmesophylla

Colourlessmesophylla

Bundlesheatha

Waterstoragea

T. pergranulata 0.6560.04 A 1.7160.04 Aa 0.0860.01 Ab – – 0.1960.01 AcT. indica subsp. indica 2.1860.07 B 3.0760.18 Ba 0.0760.01 Ab 0.7360.06 Ac 0.7560.11 Ac 0.7560.03 BcT. indica subsp. bidens 0.7460.01 A 1.1460.08 Aa 0.0860.01 Ab 0.3460.02 Bc 0.5260.02 Bd 0.2660.01 Ce

a The thickness of two adjacent cell walls was measured and divided by 2. The average number of partial cell profiles/sections examined was 29.




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Very often, two neighbouring mesophyll CWs are notvery tightly appressed to each other, having the inter-cellular space filled with fibrillar material (Fig. 4C).Plasmodesmata are more often found in the tangential(periclinal) CW between two MCs rather than in the radial(anticlinal) CW; but, in both cases, they are located in thelocal thickening of the CW (Fig. 4D). All WS cells areinterconnected by plasmodesmata, which are also locatedin a thickened area of the CW (not shown).The ultrastructure of palisade MCs and Kranz BSCs in

both subspecies of T. indica is similar in general features.

The chloroplast size (based on length) in the chlorophyl-lous and colourless MCs and BSCs is ;4–6 lm, withlittle to no difference in size between the cell types, andfrom that in MCs of T. pergranulata. The thylakoidsystem in the mesophyll chloroplasts consists of sparsegrana which have short thylakoids with a high degreeof stacking and numerous, long intergranal thylakoids(Fig. 4G, subsp. indica). Mesophyll mitochondria in thetwo subspecies are rather small (;0.4 lm) and compara-ble in size with mitochondria in MCs of T. pergranulata(Table 2). MCs usually are packed rather tightly on their

Fig. 4. Electron microscopy of cuticle, CWs, chloroplasts, and mitochondria in chlorenchyma cells in stems of T. pergranulata (A–D) and T. indicasubsp. indica (E–I). (A, E) Cuticle. (B) Mesophyll chloroplast. (C, D) Contact of two neighbouring MC walls (C) with thickened area withplasmodesmata (D). (F, G) General view of an MC (F) and mesophyll chloroplast with reduced grana (G). (H) Colourless MCs with thickened CWsand starch-accumulating chloroplasts. (I) Comparison of chloroplast structure in colourless MCs (to the left) and chlorenchyma MC chloroplasts (tothe right). (J) Structure of a chloroplast in colourless MCs. (K) Positioning of organelles in BSCs. (L) Organelles in BSCs: numerous specializedmitochondria between granal chloroplasts. (M) Colourless BSCs with starch-accumulating chloroplasts. (N, O) Plasmodesmata in the inner BS CWtowards the vascular bundle parenchyma (N) and WS tissue (O). (P) Undulating positioning of cellulose microfibrils in a thickened CW betweencolourless MCs and BSCs. (Q) Positioning of a vascular bundle facing the phloem side towards the BSC. C, cuticle; BS, bundle sheath cell; D,dendrites (polysaccharide microfibrils); M, mesophyll cell; PD, plasmodesmata; PF, pit field; Ph, phloem; X, xylem. Scale bars: 0.2 lm for A and E;0.5 lm for B–D and P; 1 lm for G, I, J, L and O; 5 lm for F and N; 10 lm for H, K, M, and Q.




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proximal end in T. indica subsp. indica, while subsp.bidens often has small air spaces between MCs and BSCs.As in the C3 T. pergranulata, MCs in both subspecieshave rather thin CWs, ;0.07–0.08 lm (Table 1), whilethe CW thickness in BSCs is 10 times higher for subsp.indica and ;7 times higher for subsp. bidens. ColourlessMCs have obviously thicker CWs than chlorenchymaMCs (Fig. 4H, I), up to 10 times in subsp. indica and fourtimes in subsp. bidens (see Table 1). The colourless MCshave plasmodesmata connections with neighbouring cells:MCs, other colourless MCs, BSCs, and colourless BSCs.These cells contain a few chloroplasts which are filledwith starch (Fig. 4I, J), and small mitochondria which arecomparable in size and internal structure with mesophyllmitochondria (the size of mitochondria in both MCs andcolourless MCs is ;0.4 lm, Table 2). The Kranz cellshave preferentially centrifugally arranged chloroplasts inboth subspecies (Figs 2G, 4K), with numerous, well-developed irregular grana interconnected by intergranalthylakoids (Fig. 4L). Specialized mitochondria are locatedbetween chloroplasts in the distal part of the cell or alongthe thinner cytoplasmic layer in the inner (proximal) partof the BSC (Fig. 4L). The BS mitochondria are ;50%larger than the MC mitochondria in both subspecies(Table 2), and they have mostly tubular cristae, with onlysome of them having a lamelliform appearance (Fig. 4L).Thick BS CWs are penetrated with pit fields on the borderwith MCs and between neighbouring BSCs, and alsoin the inner tangential CW between BSCs and WS cells(Fig. 4N, O). Colourless cells in the BS layer are similarto the chlorenchymatous Kranz BSCs in having thickCWs and pit fields with plasmodesmata. They differ fromthe Kranz cells by containing only a few chloroplasts withlarge starch grains and sparse mitochondria in a rather thincytoplasmic layer (Fig. 4M). Colourless BSCs appear tobe located internal to groups of colourless MCs. In thetwo subspecies, the CW of WS tissue is also much thickerthan that of the chlorophyllous MCs (Table 1). In subsp.indica, the thickness of the CW of WS cells is comparablewith that of BSCs and colourless mesophyll CWs, whilein subsp. bidens, the WS CWs are about half as thick as

BS CWs and similar in thickness to CWs of colourlessMCs (Table 1). The thick CWs in WS parenchyma,colourless MCs and BSCs have a similar undulateddistribution of cellulose microfibrils (Fig. 4P), which isnot observed in other tissues. In WS tissue, the cells areinterconnected with plasmodesmata, which are located ina thickened area of the CW (not shown, but similar to thatin Fig. 4D). As noted earlier, the small peripheral bundlesare often directly adjacent to BSCs, and one of the mostinteresting features of this genus is that the phloem side ofthe bundles is facing chlorenchyma tissue (Fig. 4Q).

Western blots

SDS–PAGE blots of total proteins extracted from leaveswere probed immunologically to test for C4 enzymes andRubisco LSU (Fig. 5). The molecular masses of theimmunoreactive bands corresponded to the expected massof the different polypeptides. The results show a strong

Table 2. Size of mitochondria and chloroplasts in Tecticornia species (lm)

Analysis was by one-way ANOVA with Tukey’s HSD. Means followed by a different lower-case letter within a row indicate a significant differencebetween cell types; comparison was made independently for chloroplast and mitochondria sizes (P <0.05). Means followed by a different upper-caseletter within a column indicate a significant difference between species (P <0.05). The average number of organelle sections examined in each casewas 35 for chloroplasts and 20 for mitochondria. M, mesophyll, BS, bundle sheath.

Species Chloroplast length Mitochondria small diameter

Chlorophyllous M Colourless M BS Chlorophyllous M Colourless M BS Colourless BS

T. pergranulata 4.9860.12 A – – 0.3660.03 A – – –T. indica subsp. indica 5.0560.18 Aa 5.0860.29 Aa 5.8260.21 Aa 0.3860.02 Aa 0.3960.02 Aa 0.5760.02 Ab 0.4160.01 AaT. indica subsp. bidens 6.2360.15 Ba 3.9460.31 Bb 5.2260.11 Ac 0.4060.02 Aa 0.3260.02 Aa 0.5860.02 Ab 0.6360.01 Bc

Fig. 5. Western blots for C4 enzymes and Rubisco from total proteinsextracted from green shoots of T. pergranulata, and T. indica subsp.indica and subsp. bidens. Blots were probed with antibodies raisedagainst PEPC, PPDK, NAD-ME, NADP-ME, and Rubisco, respec-tively. Numbers on the left indicate the molecular mass in kilodaltons.Western blots were replicated a minimum of three times with eachantibody with similar results.




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immunoreactive band for Rubisco LSU at 56 kDa in allspecies. Strong immunoreactivity was observed for PEPCand PPDK in the two C4 subspecies. With antibodies toC4 acid decarboxylases, there was immunolabelling forNAD-ME (65 kDa) in both subspecies of T. indica, withextremely low labelling for NADP-ME (62 kDa) (Fig. 5),and no labelling for PEP-CK in any of the species (notshown). In the C3 species T. pergranulata, there werevery low immunoreactive bands for all C4 enzymes, i.e.PEPC, PPDK, NAD-ME, and NADP-ME (Fig. 5), and nolabelling for PEP-CK (not shown).

Immunolocalization of enzymes and starch distribution

In the C3 species T. pergranulata, immunolabelling forRubisco occurs in chloroplasts of all chlorenchymacells (Fig. 6A), similar to the distribution of starch grains(Fig. 2D). The distribution of in situ immunolabelling forseveral photosynthetic enzymes in the C4 T. indica subsp.indica is shown at light microscopy (Fig. 6B, C) andelectron microscopy levels (Fig. 7) (also see Table 3 forcomparison of the density of labelling for differentphotosynthetic enzymes in different cell types for the twosubspecies). There was strong labelling for Rubisco LSUin chloroplasts in BSCs (Fig. 6B, Table 3) which alsostore starch (Fig. 2K), and the few chloroplasts found inWS cells also show labelling for Rubisco (not shown).Labelling for PEPC is high in MCs (Fig. 6C) and isconfined to the mesophyll cytosol (Fig. 7C, Table 3).Transmission electron microscopy studies of the twosubspecies of T. indica show immunolabelling forNAD-ME and GDC in BSC mitochondria (Fig. 7B).To study the possible function of colourless MCs,

immunolabelling was performed at the electron micros-copy level (see results Table 3). For both T. indicasubspecies, the labelling for PEPC is highest in thecytosol of MCs (Fig. 7C, Table 3); however, the colour-less MC also showed substantial labelling (thoughsignificantly lower than in MCs) (Fig. 7D, Table 3).Labelling for PPDK was shown to be localized pre-dominantly in chloroplasts of MCs, with lower, butsignificant, labelling in chloroplasts of colourless MCs.

Labelling for PPDK in BS chloroplasts and for PEPC inBS cytosol was similar to background (Table 3). Starchdistribution, in general, followed the pattern of Rubiscolocalization, with higher starch content in BSCs incomparison with MCs, but the largest starch granulesare localized in the colourless MCs and colourless BSCsin T. indica subsp. indica (Fig. 4H, J, M).

Gas exchange measurements, carbon isotopecomposition, and titratable acidity

Similar responses of photosynthesis to varying light wereobserved for the C3 plant T. pergranulata and C4 speciesT. indica subsp. indica and subsp. bidens. In all three taxa,photosynthesis saturates at relatively high light intensity,;1200 PPFD, but T. pergranulata reaches a highermaximum rate than the C4 species (Fig. 8). The rate ofphotosynthetic CO2 fixation was measured at varying

Fig. 6. Reflected/transmitted confocal imaging of in situ immunolocalization of photosynthetic enzymes in stems of T. pergranulata (A) and T.indica subsp. indica (B, C). Immunolabel appears as yellow dots. (A, B) Rubisco. (C) PEPC. Scale bars: 50 lm.

Fig. 7. Electron microscopy of in situ immunolocalization of PEPC,NAD-ME, and GDC in chlorenchyma cells of T. indica subsp. indica.(A) GDC and (B) NAD-ME in BSC mitochondria. Scale bars: 0.5 lm;(C, D) PEPC in chlorenchyma MC cytosol (C) and colourless MCcytosol (D).




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intercellular levels of CO2 (Ci) under atmospheric (21%)and low (2%) concentrations of O2. Under varying CO2

and ambient O2, the C3 species T. pergranulata has lowercarboxylation efficiency, and increasing rates of CO2

fixation up to a Ci of 900 lmol mol�1 (Fig. 9A), whereasthe two Kranz-type subspecies show a similar, rela-tively rapid increase in photosynthesis with increasingCi up to ;600 lmol mol�1 (Fig. 9B, C). A higher levelof O2 was inhibitory for photosynthesis rates undervarying CO2 in T. pergranulata (Fig. 9A), while bothT. indica subspecies had no inhibition of photosynthesisby O2 (Fig. 9B, C). The C* was determined for thethree taxa (Table 4) by analysis of the intercept ofCO2 response curves at different light intensities (asillustrated in Fig. 9D for T. indica subsp. bidens).C* is much lower in the Kranz-type C4 species than inthe C3 species. Both T. indica subspecies have C4-type d13C values (subsp. indica –13.7&, and subsp.bidens –15.2&), while T. pergranulata has C3-typevalues (–31.4&). Titratable acidity tests did not revealany changes in pH of cell sap during the diurnal cycle(Table 4).

Discussion

There has been a strong interest in the evolution of C4

photosynthesis in the family Chenopodiaceae, due to itsunusually high diversity, with different Kranz and non-Kranz C4 leaf types as well as variation in C3 leaf types(Monteil, 1906; Carolin et al., 1975; Fisher et al., 1997;Jacobs, 2001; Pyankov et al., 2001a, b; Schutze et al.,2003; Kapralov et al., 2006). Halosarcia, as traditionallydefined, has been shown to be paraphyletic in relation toother Australian Chenopodiaceae genera (Shepherd et al.,2004; Kadereit et al., 2006; this study). The monophylyobtained from molecular studies has been supportedbased on morphological characters which show a high

level of homoplasy (Shepherd et al., 2005; Shepherd andWilson, 2007). Because of this, these genera have allrecently been reorganized into a more broadly definedTecticornia (Shepherd and Wilson, 2007), which isaccepted here. While this clade of species is predomi-nantly Australian in distribution, it is also found on othercontinents, including southern Asia (Malaysia, Sri Lanka,India, and Pakistan) and tropical East Africa along coastaland inland saline areas. Interestingly, the only speciespreviously described as having C4 photosynthesis, T.indica, is also one of the few Australian chenopodlineages also to be found outside of the continent. Carolinet al. (1982) identified four subspecies of Halosarcia (¼Tecticornia) indica (bidens, indica, julacea, and leiosta-chya) as C4 plants. From molecular phylogeny based onnuclear DNA internal transcribed spacer (ITS) data,T. indica and most of its subspecies form a stronglysupported clade with undescribed entities previously

Table 3. Cellular immunogold labelling of photosynthetic enzymes in various cells of stems of Tecticornia indica subsp. indica andsubsp. bidens (number of gold particles per 1 lm2)

Analysis was by one-way ANOVA with Tukey’s HSD. Means followed by a different lower-case letter within a row indicate a significant differencebetween cell types (P <0.05). For PEPC comparisons were made between numbers of particles in the cytosol of three cell types, for PPDK andRubisco comparisons were made between the numbers of particles in chloroplasts of three types of cells. The number of gold particles is given as themean 6SE. The average number of partial cell profiles/sections examined was 20.

Species Chlorophyllous mesophyll cells Colourless mesophyll cells Bundle sheath cells

PEPC Organelles Cyt Background Organelles Cyt Background Organelles Cyt BackgroundT. indica subsp. indica 1.360.4 7.361.1 a 0.660.2 0.960.3 3.160.8 b 1.960.5 1.060.4 1.360.6 c 0.860.4T. indica subsp. bidens 1.560.4 9.461.1 a 1.060.3 2.660.7 6.361.1 b 1.260.3 1.160.2 1.560.3 c 0.860.2

PPDK A B C A B C A B CT. indica subsp. indica 9.661.0 a 1.460.2 0.560.2 4.860.8 b 0.960.3 1.960.5 2.760.4 c 0.960.1 2.861.1T. indica subsp. bidens 15.061.2 a 1.760.3 1.760.2 5.861.0 b 0.560.4 1.960.6 1.560.3 b 0.760.3 2.061.1

Rubisco A B C A B C A B CT. indica subsp. indica 2.860.4 a 0.0660.03 0.360.1 4.760.7 a 0.260.1 0.760.2 20.861.3 b 0.260.1 0.460.03T. indica subsp. bidens 4.160.5 a 0.3560.2 0.460.3 11.361.0 b 0.460.2 0.460.2 22.862.1 c 0.260.1 0.460.2

A, Chloroplast; B, Cyt + other organelles; C, Background.

Fig. 8. Rates of CO2 fixation in response to varying light at 25 �C and370 lmol mol�1 of CO2 in T. pergranulata, and T. indica subsp. indicaand subsp. bidens. The results represent the average of three replicationsfrom measurements made on different branches.




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referred to as ‘Yanneri Lake’ (Shepherd and Wilson,2007), which has been suggested also to be C4 (KShepherd, personal communication). This may indicatea single origin of C4 photosynthesis in Salicornioideae.However, T. indica subsp. julacea is not part of the T.indica clade in the ITS tree (Kadereit et al., 2006), only inthe chloroplast DNA trnL tree of Shepherd et al. (2004).The phylogenetic positions of the C3 and C4 taxa utilizedin this study were verified by comparison with some otherspecies in genus Tecticornia and related genera insubfamily Salicornioideae using ITS as a marker andmaximum likelihood analysis (Fig. S1 and Table S1 inSupplementary material available at JXB online). Obvi-ously, a more detailed analysis of T. indica subspecies is

needed. The differences between two subspecies of T.indica, subsp. indica and subsp. bidens, including theirdifferent habit (subsp. indica is a prostrate dwarf shrub upto 50 cm, rarely to 1 m, versus subsp. bidens which is anerect shrub up to 2 m tall) (Wilson, 1980), macroscopic,microscopic, and genetic differences described in thispaper, and their different geography, collectively supportspecifically different entities. This needs to be clarified inthe re-evaluation of the taxonomic status of this complexin a broad geographical context. These results, anda similar case already discussed in the genus Bienertia(Akhani et al., 2005), indicate that the taxonomy ofseveral critical groups of Chenopodiaceae needs to bereassessed using multidisciplinary approaches.

Table 4. CO2 compensation point, carbon isotope discrimination (d13C), and test for CAM in Tecticornia species

For determination of C* see Fig. 9. For d13C and titratable acidity n¼2.

Species C* (lmol mol�1) d13C (&) Titratable acidity (leq g FW�1)

End of the night Middle of the day End of the day

T. pergranulata 34.2 –31.460.05 1.8260.23 1.9360.04 1.9660.08T. indica subsp. indica 5.2 –13.760.01 2.4260.57 2.9760.53 2.5060.13T. indica subsp. bidens 4.8 –15.260.01 3.9960.10 4.0160.09 3.6660.21

Fig. 9. Rates of CO2 fixation in response to varying intercellular levels of CO2 at 25� C and 900 PPFD in T. pergranulata (A), T. indica subsp.indica (B), and subsp. bidens (C). The results represent the average that was taken of ambient to low CO2 response, and ambient to high CO2

response, from separate measurements on 2–3 branches. (D) Illustration of calculation of C* from CO2 response curves at 25 �C under four lightintensities with T. indica subsp. bidens: 90 (line 1), 170 (line 2), 260 (line 3), and 360 (line 4) PPFD. Each light level is the response to tworeplications.




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Anatomical features

The structure of chlorenchyma in C3 T. pergranulata,consisting of two layers of elongated MCs around theperiphery of cylindrical leaves or aphyllous stems, israther typical for different C3 representatives of Salicorn-ioideae and Salsoloideae; in aphyllous species, smallreduced scale-like leaves have similar chlorenchymatissue only on their abaxial side. This type of structure,with a peripheral position of chlorenchyma and a networkof small vascular bundles, and a central cylinder (in stems)or main vascular bundle (in leaves) in the centre, has beencalled ‘centric’ (Metcalfe and Chalk, 1950), ‘sympeg-moid’ (Carolin et al., 1975), or ‘arco-vascular’ (Vasilev-skaya and Butnik, 1981). It has also been described inSalsoloideae as having peripheral vascular bundles withthe xylem side facing the chlorenchyma (Carolin et al.,1975; also see figs 7, 8 and 11, 12 in Pyankov et al.,1997; fig. 2C, D in Voznesenskaya et al., 2001; figs 2, 5,7 in Voznesenskaya et al., 2003). Although not perfect,this is a convenient way of identifying this anatomicaltype when it occurs, and we consider that the term‘centric’ reflects well all features of C3 anatomy in thesecases. The characteristic feature distinguishing T. pergra-nulata from C3 Salsoloideae species which have a similarstructure is the positioning of peripheral vascular bundles.In Tecticornia, the phloem side of the small peripheralbundles faces towards the chlorenchyma tissue. Also, allperipheral bundles in C3 T. pergranulata are separatedfrom the chlorenchyma tissue by one layer of large WScells, while in C3 or C3–C4 Salsola species, the peripheralvascular bundles are separated from chlorenchyma cellsby rather small parenchyma cells representing parenchy-matous BS around peripheral vascular bundles (Pyankovet al., 1997; Voznesenskaya et al., 2001; Akhani andGhasemkhani, 2007; EV Voznesenskaya, unpublishedresults). In other species of the Australian Salicornioideaesuch as Tecticornia s. str. and Pachycornia and Sarcocor-nia, many of the vascular bundles are adjacent tochlorenchyma (Carolin et al., 1982). While species in thegenus Salicornia have a similar positioning of the phloemto that in Tecticornia, the peripheral bundles are oftenadjacent to the chlorenchyma tissue (personal observationof EV Voznesenskaya and NK Koteyeva, unpublishedresults). The structure and position of peripheral vascularbundles in C3 T. pergranulata represent a rather distinc-tive feature, which may also be characteristic for someother Salicornioideae. Accepting this type of anatomy as C3

centric, as a minimum two variants should be mentionedaccording to the positioning of peripheral vascular bundles.In the two C4 subspecies of T. indica, chlorenchyma

tissue consists of two cell layers, elongated MCs androundish BSCs, on the periphery of the stems andrudimentary leaves. The present observation of parader-mal sections revealed that the islands of chlorenchyma

cells are surrounded by sections of large, colourless MCswith thick CWs, which consist of one to three cells across,as previously reported (Carolin et al., 1982). Kadereitet al. (2003) distinguished the anatomical type in T. indicaas Kranz-halosarcoid, based on the presence of colourlessMCs and the centrifugal position of chloroplasts in BSCs.An additional distinguishing anatomical feature of thisKranz type is the position of peripheral vascular bundlesdirectly adjacent to BSCs, with the phloem facing thechlorenchyma tissue, as was also observed in C3

T. pergranulata. This differs from salsoloid-type C4

species, where the xylem in the peripheral vascularbundles faces the chlorenchyma tissue (see Olesen, 1974;Voznesenskaya, 1976a, b; figs 7, 8, 11, 12 in Pyankovet al., 1997; fig. 2A, B in Voznesenskaya et al., 2001;figs 2, 5, 7 in Voznesenskaya et al., 2003). Interestingly,a similar positioning of peripheral vascular bundles, withtheir phloem side towards the chlorenchyma, was onlypreviously mentioned in the ‘single-cell functioning’ C4

species, Suaeda (¼Borszczowia) aralocaspica (Freitagand Stichler, 2000). It was thought (Olesen, 1974) thatsuch positioning of vascular bundles could facilitate thetransport of assimilates and water, but this idea needsfurther investigation. Thus, the type of chlorenchymastructure in C4 Tecticornia represents a unique variationof Kranz anatomy with discontinuous chlorenchyma,interrupted by the thick-walled colourless cells in bothlayers, mesophyll and BS, centrifugally arranged organ-elles in BSCs, and positioning of peripheral vascularbundles with their phloem side to the chlorenchyma. Thistype of anatomy was designated Kranz-halosarcoid byKadereit et al. (2003) according to the previous name ofthe genus, which is now changed to Kranz-tecticornoidtype. In general, this type of anatomy can be described asKranz centric discontinuous with the specific position ofvascular bundles and chloroplasts in BSCs.According to Carolin et al. (1982), Fahn and Arzee

(1959), and Al-Turki et al. (2003), in all species ofsubfamily Salicorniodeae studied, the network of vascularbundles in the fleshy cortex is derived from the leafbundle of the upper internode. The type of venation inreduced leaves and cortex was classified as Salicornia-Arthrocnemum type (Fahn and Arzee, 1959), where thedescending network venation of the cortex was derivedonly from lateral branches of the leaf strands. The study ofvenation in C3 and C4 Tecticornia species also showedthis type of venation and origin of peripheral bundles,with certain differences in the structure of the primaryvascular system between T. pergranulata and T. indicasubsp. bidens; in general, the Salicornia-Arthrocnemumtype of venation was suggested to be more advanced incomparison with venation in Kochia-Bassia or Rhagodia-Atriplex types (Bisalputra, 1962). There has been anextensive discussion of the origin of the fleshy cortex inarticulated Chenopodiaceae species (see Fahn and Arzee,




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1959). From examination of the origin of the cortexduring plant development, it was concluded that theassimilating cortex is not a product of leaf fusion andadnation to the stem, but rather is a result of simultaneousgrowth of the leaf basis and cortex. It was shown that thedevelopment of the reduced leaves in species with suchshoots is similar to that of ordinary foliar leaves(Vasilevskaya, 1955; Werker and Fahn, 1966) and thatthe fleshy tissue external to the central cylinder of theseplants develops as a result of intercalary growth at thebase of each internode and should be regarded as truecortex (Vasilevskaya, 1955; Fahn and Arzee, 1959). Therelationship between positioning of the small peripheralbundles and transport of assimilates in different Chenopod-iaceae species needs additional study.It is interesting to note that chenopods with a fleshy

cortex have a special form of secondary growth andperiderm formation which was previously studied in thegenus Haloxylon. The secondary cambium, as well as theperidermal cambium, originates in the pericycle, which isinternal to the endoderm, and usually after formation ofthe periderm the outer fleshy chlorophyllous cortexwithers and dries up (Arcihovskii, 1928; Vosnesenskayaand Steshenko, 1974). While the process of secondarygrowth was not studied in Tecticornia, light microscopyimages show very similar secondary growth to that inHaloxylon.

Tecticornia indica: C4 biochemical subtype andenzyme compartmentation

The high levels of C4 cycle enzymes PPDK, PEPC, andNAD-ME in T. indica are indicative of C4 photosynthesis,as compared with the very low levels of these enzymes inthe C3 species T. pergranulata. Analysis by western blotsfor C4 acid decarboxylases shows that T. indica is anNAD-ME-type C4 species. Generally, consistent resultshave been obtained in subtyping C4 species by immuno-detection versus enzymatic assay of C4 decarboxylases(Walker et al., 1997; Wingler et al., 1999; Pyankov et al.,2000; Voznesenskaya et al., 2002). Immunolocalizationstudies show selective compartmentation of PPDK andPEPC in MCs, and Rubisco in BSCs, in the twosubspecies of T. indica, characteristic of C4 plants. Highlevels of starch accumulate in the BSC chloroplastscompared with MC chloroplasts (see also Carolin et al.,1982). Also, NAD-ME and GDC are selectively localizedin mitochondria of BSCs, as expected for NAD-ME-typeC4 species.

Ultrastructural features of photosynthetic tissue

The ultrastructural characteristics of chlorenchyma cells inT. pergranulata are typical of other C3 species, withchloroplasts and mitochondria around the periphery ofMCs. In the two C4 subspecies of T. indica, there is

differentiation of chloroplasts and mitochondria betweenMCs and BSCs. There are numerous mitochondria inBSCs which, along with chloroplasts, are predominantlylocated in the centrifugal position, as was also observedby Carolin et al. (1982) and Jacobs (2001). Themitochondria in BSCs are ;50% larger than in MCs,while the chloroplasts in the two cell types are similarin size. However, the mesophyll chloroplasts have areduction of grana with prevalence of intergranal thylak-oids compared with BS chloroplasts. The abundance ofmitochondria in BSCs and the reduction of grana inmesophyll compared with BS chloroplasts are typicalof NAD-ME-type C4 species (Carolin et al., 1975;Voznesenskaya, 1976a, b; Gamaley, 1985; Voznesenskayaand Gamaley, 1986; Fisher et al., 1997).In most NAD-ME-type C4 species, including dicots and

monocots, the chloroplasts are in a centripetal position(Gutierrez et al., 1974; Hattersley, 1987; Dengler andNelson, 1999). However, there are established cases ofNAD-ME-type species having BS chloroplasts in thecentrifugal position. With respect to dicots, the centrifugalposition of chloroplasts in BSCs of T. indica is similar tothat found in Suaeda species having a schoberia leaf typewith NAD-ME C4 photosynthesis (Schutze et al., 2003;Voznesenskaya et al., 2007). This also occurs in Trian-thema triquetra, family Aizoaceae, which has atriplicoidleaf anatomy and an unspecified biochemical subtype, butwith ultrastructural features characteristic of NAD-MEspecies (Carolin et al., 1978). With respect to monocots,the centrifugal position of BS chloroplasts has been foundin several NAD-ME-type species: in spp. of Panicum sect.Dichotomiflora, in Eragrostis, and in Enneapogon (Ohsugiet al., 1982; Hattersley, 1987). Whether there is functionalsignificance to the chloroplast position, or whether it isonly indicative of alternative forms of C4, is unknown.Many C4 species have BSCs with thickened CWs (see

Sage and Monson, 1999). Among C4 NAD-ME species infamily Chenopodiaceae, it is possible to distinguish twogroups according to the thickness of their BS CWs. MostAtriplex and Suaeda species have rather thin CWs, whilerepresentatives studied from tribe Caroxyloneae withNAD-ME-type anatomy, Climacoptera transoxana,Halocharis hispida, and Salsola rigida (¼Caroxylonorientale) (Akhani et al., 2007), have very thick BS CWs(Voznesenskaya, 1976b), similar to those in C4 Tecticor-nia. The most interesting feature of BSC structure in theC4 T. indica subspecies is the presence of intercellularconnections by plasmodesmata, not only in the outertangential CW (between BSCs and MCs), but also in theinner tangential CW, between BSCs and WS tissue, andbetween BSCs and vascular bundle parenchyma cells.This feature suggests symplastic transport of assimilatesfrom chlorenchyma to the vascular tissue in these C4

species. Also, in T. indica, the WS cells have rather thickCWs which are interconnected by plasmodesmata.




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Fluorescence of chloroplasts and cell walls, andlignification

When excited by UV radiation, leaves of all plants haveintensive red fluorescence from all chlorophyll-containingcells. Obviously, the most intensive red colour in sectionsof stems was in the chlorenchyma tissue in the outer cortexlayers, with lower red fluorescence from several otherchloroplast-containing parenchymatous tissues includingthe pith, xylem, and phloem parenchyma, and the perider-mal parenchyma (the phelloderm) which is located justoutside the central cylinder. Possible functions of theinternal chlorophyll-containing tissues were studied in someSalicornioideae species by Redondo-Gomez et al. (2005).Certain groups of green plants exhibit a genuine blue

fluorescence from their CW due to accumulation ofphenolic substances, especially lignins and/or suberins. Instem sections of the Tecticornia taxa studied, the brightestblue fluorescence was emitted from lignified fibres,sclerenchyma, and xylem elements, and from the suber-ized layers outside the central cylinder representing theperiderm. The blue fluorescence of non-lignified CWs (i.e.those that have a negative phloroglucinol-HCl test)changes to green with increasing intensity after treatmentwith 0.1 M NH4OH, indicating the presence of boundferulic acid (Rudall and Caddick, 1994). According toprevious studies, families of monocotyledons can bedivided into two groups depending on the UV fluores-cence behaviour of their CW and presence or absence ofbound ferulic acid (Harris and Hartley, 1976; Harris andHartley, 1980). In dicots, wall-bound ferulic acid has onlybeen found in the order Caryophyllales, and was pre-viously shown for eight species of family Chenopodiaceae(Hartley and Harris, 1981).In T. pergranulata and T. indica subspecies, the non-

lignified CWs of assimilating organs fluoresce blue underUV radiation and change colour to intense green afterNH4OH treatment, indicating the presence of CW-boundferulic acid. Intense fluorescence following NH4OHtreatment was found in all three representatives and, thus,it does not depend on photosynthetic type. There wasdifferential distribution of fluorescence intensity in differ-ent tissues, with maximum green fluorescence afterNH4OH treatment in epidermal and WS tissues. In bothC4 subspecies, the walls of BSCs fluoresce more intensegreen than the chlorenchymatous MCs, with the highestintensity in the thick-walled, colourless MCs. In C3 andC4 Tecticornia, the intensity of green fluorescencefollowing NH4OH treatment tended to correspond to CWthickness. The epidermis has very thick CWs, and the WStissue has much thicker CWs than the MCs in bothspecies. In the C4 subspecies of T. indica, the WS tissue,BSCs, and colourless MCs have higher fluorescence andmuch thicker CWs than the MCs. Carolin et al. (1975)mentioned different staining of the mesophyll and BS CW

by electron microscopy; however, no differences wereobserved in the present study. Nevertheless, most of thethickened CWs in T. indica, including BSCs, colourlessMCs, and WS tissue, have a specific undulating distributionof cellulose microfibrils which is absent in all other tissues.With respect to the possible functions of CW ferulic

acid, it has been suggested that, in certain groups of plants(in particular in Poaceae), ferulic acid in the walls ofepidermal cells absorbs UV-B radiation and protects thephotosynthetic apparatus (Lichtenthaler and Schweiger,1998). Wakabayashi et al. (1997) showed that increasedlevels of ferulic acid led to decreased CW extensibilityand to significantly increased mechanical strength oftissues. In some desert plants (e.g. Tecticornia), in whichthe stem is the main carbon-assimilating organ, there islittle tissue to give mechanical support; thus, the presenceof ferulic acid may provide strength to the CW to supportthe stems. It has been shown that the quantity of ferulicacid increases under water and osmotic stresses, whichwas suggested to facilitate adaptation to dry and salineenvironments (Wakabayashi et al., 1997; Fan et al., 2006).For function of C4 photosynthesis, there needs to be

resistance to loss of CO2 from sites of C4 acid de-carboxylation in BSCs, in order for it to be assimilatedeffectively by Rubisco, and a number of factors contributeto this to varying degrees, depending on the species andC4 subtype (von Caemmerer and Furbank, 2003). The BSCWs may contribute to this resistance, depending onthickness and composition, and it has long been recog-nized that BSCs in some C4 species have a suberizedlamella, which is thought to contribute to diffusiveresistance. In T. indica, fluorescence and histochemicalanalyses indicate that BSCs lack lignin and suberization,and that the higher apparent content of ferulic acid inBSCs corresponds to a thicker CW. In the two C4

subspecies, the BS CW is 7- to 10-fold thicker than theMC CW; thus, the thicker CW may contribute to theresistance to leakage of CO2 from BSCs. The chloroplastsin BSCs of T. indica are predominantly located ina centrifugal position, which would reduce diffusiveresistance through the liquid phase, and increase potentialfor leakage from sites of decarboxylation to the exterior ofthe cell. However, the mitochondria in BSCs, which arethe site of C4 acid decarboxylation via NAD-ME, arepositioned internal to the chloroplasts, which is favourablefor refixation of CO2 by Rubisco in the BS chloroplasts.

Photosynthetic CO2 exchange, carbon isotopecomposition, and titratable acidity

The two C4 subspecies of T. indica have C4 type d13Cvalues (subsp. indica –13.7& and subsp. bidens –15.2&)and low C* values (subsp. indica 5.2 and subsp bidens4.8 lmol CO2 mol�1, Table 4) which indicates theefficiency of function of C4, while T. pergranulata has C3




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type C* (34.2 lmol mol�1) and C3 type d13C values(–31.4&). The d13C values of these species are consistentwith earlier results of Carolin et al. (1982), who obtainedvalues of –12.2& to – 14.2&. The CO2 response curvesunder 2% versus 21% O2 show that CO2 assimilation in T.indica is insensitive to O2, which is characteristic of C4

plants. CO2 assimilation in T. pergranulata is inhibited by21% O2 under limiting CO2 due to photorespiration andlack of a CO2-concentrating mechanism. The results showthe C4 Tecticornia would have an advantage underconditions where CO2 is limiting. C3 plants often havea lower light saturation of photosynthesis than C4 plants.However, the light response curves were similar for the C3

and C4 Tecticornia species, which may be related to thethick stems requiring high light to saturate photosynthesis.The absence of nocturnal acidification of cell sap in all

three representatives of this genus indicates that they donot have the CAM type of photosynthesis.

Possible functions of unique colourless cells

In the stem tissue of C4 T. indica, there is a wreath ofphotosynthetic tissue near the periphery. However, this isinterrupted by an unusual co-occurrence of colourlessMCs and BSCs within the layers of chlorenchyma,characteristic of Kranz anatomy. The area of the colour-less MCs in the longitudinal plane appears to be greaterthan that of the colourless BSCs. The very few plastidswhich occur in these cells have high levels of starch,although the Kranz BSCs are the main sites of starchstorage. Analysis of the enzyme composition of thecolourless MCs also showed they did not have meso-phyll-type specialization for C4 photosynthesis. ColourlessMCs were not observed in the C3 species T. pergranulata,which raises the question as to whether this feature mayhave co-evolved with evolution of C4 photosynthesis inthe genus.There has been speculation as to how windows in some

succulent species may influence photosynthesis (seeEgbert and Martin, 2002). One possible function of thesecolourless areas within Kranz anatomy is to distributesome of the incident radiation on the tissue inside thestem. As direct sunlight is received from one side of thestem, the colourless areas may increase penetration oflight to the opposite side, which could increase efficiencyof photosynthesis in densely growing shoots. Recently, itwas noted that windows may influence photosynthesis insome plants by illuminating the chlorenchyma from twosides, inside and outside; also, in some CAM species,there is evidence that windows increase infrared radiationinside the tissue, possibly functioning to optimize leaftemperature (C Martin, personal communication).Another function may be mechanical, contributing to

stem strength, since the multicellular network of colour-less MCs have much thicker CWs than the MCs. Similar

structures have been observed in many xerophytic species,where the patches of green mesophyll are interrupted bycolourless cells, which can occur as fibre strands (inorders Fabales and Asterales), by modified thick-walledchlorenchyma or parenchyma sheath cells, elongatedperpendicular to the surface (in Restionaceae) (B}ocherand Lyshede, 1972; Fahn and Cutler, 1992), or byseparate fibres or tracheids in Arthrocnemum and Salicor-nia (Chenopodiaceae) (SaadEddin and Doddema, 1986;Fahn and Cutler, 1992; Keshavarzi and Zare, 2006). Thistype of structure was thought to have a supportingfunction, preventing collapse of soft chlorenchyma tissueduring water stress, or this compartmentation may helpprevent spread of fungal infection from one patch ofchlorenchyma to others.There is also a distinctive colourless region of cells

at the tips of the reduced leaves in both the C3

T. pergranulata and C4 T. indica subspecies. This featuremay increase penetration of light into the photosyntheticcortex of the tissue. Also, in Suaeda monoica, twotranslucent gaps have been observed at the edges of leaves(Shomer-Ilan et al., 1975; Schutze et al., 2003).

Conclusions

Family Chenopodiaceace has many C4 species occurringin three subfamilies, Chenopodioideae, Salsoloideae, andSuaedoideae. However, in species of subfamily Salicorn-ioideae, which have stems as the major photosyntheticorgan, Tecticornia indica s. l. and an undescribed taxonTecticornia ‘Yanneri Lake’ form a single well-supportedclade which appear to be the only C4 lineages in thesubfamily (this study; K Shepherd, personal communica-tion; Carolin et al., 1982). Tecticornia indica has anunusual type of Kranz anatomy with a network ofcolourless MCs surrounding the patches of MCs withinthe outer layer of chlorenchyma. These colourless cells,which have thick CWs and a few chloroplasts with limiteddevelopment for photosynthesis, may function to givea more optimum distribution of incident radiation in thephotosynthetic tissue. Tecticornia indica is an NAD-MEC4 plant having chloroplast structural features, andabundance of mitochondria in BSCs, typical of this C4

subgroup. C4-type d13C values, low C*, and O2 in-sensitivity of carbon assimilation indicate effective func-tion of C4 photosynthesis. The positioning of themitochondria, which is the site of C4 acid decarboxyl-ation, internal to the centrifugally located BS chloroplasts,and the thickened BS CWs may support efficient donationof CO2 to Rubisco. This study describes a unique C4

structural type of anatomy, Kranz-tecticornoid, in thegenus Tecticornia. Further research is needed on Tecticor-nia species and subspecies to determine if there is morediversity in forms of photosynthesis in the genus (i.e.other C4 species, or C3–C4 intermediates), and to




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determine through structural and phylogenetic studies howC4 may have evolved in this subfamily.

Supplementary material

The Supplementary material available at JXB onlineconsists of one figure and one table. They show thephylogenetic position within Tecticornia of taxa analysedin this study based on ITS sequence data. Table S1 liststhe taxa sequenced and Fig. S1 shows a phylogram.

Acknowledgements

This material is based upon work supported by the National ScienceFoundation under Grant nos IBN-0236959 and IBN-0641232, byCivilian Research and Development Foundation grants RB1-2502-ST-03 and RUB1-2829-ST-06, and by Russian Foundation of BasicResearch grant 05-04-49622. HA acknowledges sabbatical andresearch support from the Research Council, University of Tehran(Project No. 6104037/1/01), and travel support from the IslamicDevelopment Bank; also the help of Drs A Khan and S Gulzarduring the field expedition to Pakistan. We also thank theFranceschi Microscopy and Imaging Center of Washington StateUniversity for use of their facilities and staff assistance, MGhasemkhani and M Smith for some assistance in growing plants,C Cody for plant growth management, and Dr G Barrett forproviding seeds of Australian T. pergranulata and T. indica subsp.bidens.

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